Can Timing and Method of Barley Cover Crop Termination Impact Pests and Beneficials within a Subsequent Soybean Planting?

Alan W. Leslie, Armando Rosario-Lebron, Guihua Chen and Cerruti R. R. Hooks
Department of Entomology, College of Computer, Mathematical, and Natural Sciences


This extension article is meant to serve as a condensed write-up of a completed field study. Full-text of the published work can be viewed via open access at Cover cropping has long been used as a method of reducing soil erosion, increasing soil quality, and suppressing weeds. However, impacts of cover crops in cropping systems differ and can be affected by timing and method of their termination. Field trials were conducted over two field seasons and at two sites in Maryland to examine how varying the date and method of terminating a barley (Hordeum vulgare) winter cover crop affects arthropods (insects and spiders) in succeeding no-till soybean (Glycine max) plantings. Experimental treatments included early-kill with pre- and post-emergent herbicides (EK), late-kill with pre- and post-emergent herbicides (LK), late-kill with a flail mower and pre-emergent herbicide (FM), and a fallow/bare-ground check with pre- and post-emergent herbicides (BG). Terminating barley late (i.e., just prior to soybean planting) resulted in significantly greater biomass accumulation in LK and FM than EK. However, method and timing of termination had no effect on communities of pest and beneficial arthropods in the soybean canopy. Results from this experiment suggest that terminating the cover crop early or late or using a mower or burn-down herbicide to kill the cover crop will result in similar species and number of arthropods within the soybean canopy.


Cover cropping can be a viable weed management tool in conservation agriculture systems. When cover crops are terminated in reduced- and no-till cropping systems, resulting residues that remain on the soil surface can help prevent weed establishment. Thus, it is well known that cover crop residue impacts weed populations. More specifically, some of these studies were designed to examine how method and timing of cover crop termination practices impact weed populations in grain crops. However, impacts of these practices on arthropod populations are rarely considered. Despite this, studies have shown that cover crops can impact arthropod numbers in succeeding agronomic crops. Some insect pests shown to be impacted by cover crop residue include the potato leafhopper (Empoasca fabae), bean leaf beetle (Cerotoma trifurcata), and Japanese beetle (Popillia japonica) in soybean as well as thrips in cotton (Gossypium hirsutum). In addition to insect pests, their natural enemies may be influenced by cover crop residue.

The overall goal of this study was to investigate how different cover crop termination practices impact populations of insect pests and their natural enemies within no-till soybean plantings. Specific objectives were to compare the influence of termination method (chemical versus mechanical) and timing (early versus late) on arthropod populations. Barley was chosen as the test cover crop partially because of its accessibility and popularity among producers.

Materials and Methods

Field experiments were conducted at the University of Maryland’s Central Maryland Research and Education Center at the Upper Marlboro and Beltsville farm sites in 2013 and 2014. Each field experiment consisted of four treatments, including three cover crop termination methods and a fallow/bare-ground control. The three cover crop treatments included: (1) early-kill (EK), in which the cover crop was sprayed with post- and pre-emergent herbicides in mid-April; (2) late-kill (LK), in which the cover crop was sprayed with post- and pre-emergent herbicides in late May; and (3) flail-mowed (FM), in which the cover crop was sprayed with a pre-emergent herbicide and mowed in late May. An early-kill, flail-mowed treatment was not included in the experiment because mowing typically does not kill cover crops at early stages of development and farmers do not use this method. The bare-ground treatment (BG) remained fallow after the previous crop was harvested and received the same post- and pre-emergent herbicide applications as LK.

The EK treatment was sprayed with a post- and pre-emergent herbicide mixture on 15 April at Beltsville and 16 April at Upper Marlboro in 2013 and on 18 April at both sites in 2014. The LK treatment was sprayed with a post- and pre-emergent herbicide mixture on the day soybeans were planted. The BG treatment received the same spray protocol as LK. On the day soybeans were planted, the FM treatment was sprayed with a pre-emergent herbicide and the cover crop was mowed. The soybean was planted on 21 May at Beltsville and 20 May at Upper Marlboro in 2013 and 27 May 2014 at both sites. Soybeans were planted in wide rows [76 cm (30 inch) inter-row spacing] at Beltsville and narrow rows [18 cm (7 inch) inter-row spacing] at Upper Marlboro. A late-season herbicide application was applied to all plots at the Beltsville location as a “rescue” herbicide treatment primarily for large crabgrass (Digitaria sanguinalis).

Data collection. Data on vegetative (cover crop and weed) biomass, abundance of weeds and arthropods, soil moisture as well as yield were collected during this investigation. Data on weed population, soil moisture and yield will be presented in a future edition of Agronomy News. To quantify cumulative barley and weed biomass production, cover crop and weed biomass were measured in each plot just prior to their termination. Arthropod abundances were monitored with the use of a sweep net from the R1 through R5 soybean growth stages. Arthropods collected were divided into i) natural enemies (predators – arthropods that prey on herbivores & parasitoids insects, especially wasps, that complete their development within the body of another insect eventually killing it) and ii) herbivores (insects that feed on plants). Arthropods were separated further according to seven feeding habits (guilds). The seven feeding habits that we used included 1) chewing predators, 2) sucking predators, 3) parasitoids, 4) plant-sucking herbivores, 5) pod feeders, 6) foliage feeders and 7) spiders. Though they are predators, spiders were placed into a separate predatory feeding guild.


Vegetative biomass. At each farm site, flail-mowed (FM) and late-killed (LK) treatments had higher plant biomass than early-killed (EK) or bare-ground (BG) treatments (Table 1). Total barley biomass in LK and FM treatments were more than two times greater at Beltsville than Upper Marlboro. No differences were detected in plant biomass between BG and EK treatments within each site, but there was greater weed biomass in the BG treatment at Beltsville than Upper Marlboro (Table 1).

Table 1. Cover crop and weed dry biomass just prior to their termination.

Site Treatment Mass ± SEM (kg ha−1)
Beltsville Early Kill 160.1 ± 60.5 cd1
Late Kill 2211.9 ± 83.2 a
Flail Mow 2123.4 ± 112.9 a
Bare Ground 896.0 ± 254.3 bc
Upper Marlboro Early Kill 85.8 ± 21.2 d
Late Kill 753.4 ± 100.9 b
Flail Mow 851.8 ± 62.7 b
Bare Ground 120.4 ± 27.2 d

1Different letters indicate that means are significantly different.

Arthropod Counts. In total, 54 families of arthropods were collected from sweep samples which included a total of 11,344 specimens (Table 2). Approximately 98% of arthropods collected could be assigned to one of the seven feeding guilds used. Three feeding guilds, which included plant-sucking herbivores (25%), foliage-feeding herbivores (24%), and sucking predators (21%), accounted for 70% of the entire arthropod community sampled.

The abundance of arthropods from each feeding guild was similar among treatments. However, there was a significant effect of soybean development stage on all feeding guilds. In general, parasitoid, chewing predator, and sucking predator guilds reached greatest abundance later in the season (R4 or R5 stage). In contrast, numbers of foliage feeding and plant sucking herbivores peaked earlier in the growing season at the R2 or R3 stage (Table 3). Sucking predators and spiders were found in greater numbers in Beltsville than Upper Marlboro across all soybean growth stages.

Table 2. Arthropod feeding guilds, families and their abundances. Numbers represent total abundance across all sample dates.

Beltsville Upper Marlboro
Feeding Guild Family 2013 2014 2013 2014
Spider Salticidae 25 47 24 48
Araneidae 5 69 0 37
Oxyopidae 149 101 42 94
Thomisidae 18 16 35 11
Lycosidae 0 12 0 26
Clubionidae 0 3 0 0
Ctenidae 0 1 0 0
Tetragnathidae 0 8 0 9
Linyphiidae 0 4 0 2
Pholcidae 0 1 0 0
Parasitoid Platygastridae 159 11 57 0
unspecified1 0 407 0 104
Sceleonidae 10 23 1 15
Chalcididae 0 2 0 3
Proctotrupidae 0 1 0 2
Braconidae 0 76 0 29
Eulophidae 0 18 0 5
Ichneumonidae 0 8 0 3
Tiphiidae 0 178 0 25
Aphelinidae 0 1 0 1
Encyrtidae 0 1 0 0
Mymaridae 0 0 0 1
Eurytomidae 0 0 0 1
Trichogrammatidae 0 0 0 2
Chewing predator Asilidae 5 6 0 0
Mantidae 1 1 1 0
Coccinellidae 21 262 36 193
Carabidae 0 5 0 3
Syrphidae 0 101 0 4
Cantharidae 0 0 0 1
Sucking predator Geocoridae 543 346 223 326
Pentatomidae 3 8 1 1
Chrysopidae 5 1 3 13
Anthocoridae 48 166 225 161
Nabidae 100 340 37 93
Hemerobiidae 0 10 0 3
Reduviidae 0 0 0 4
Foliage feeder Coccinellidae 287 92 43 1
Erebidae 346 756 274 455
Meloidae 0 1 0 0
Scarabaeidae 428 284 90 96
Chrysomelidae 2 345 0 254
Noctuidae 0 1 0 0
Hesperiidae 0 3 0 0
Plant sucking Cicadellidae 32 732 109 689
Membracidae 22 0 40 18
unspecified 404 0 896 0
Aphididae 0 0 0 60
Pod feeder Pentatomidae 33 164 69 115
Miridae 112 102 108 229
Unassigned unspecified 0 2 67 30
Chrysomelidae 0 0 0 2
Curculionidae 2 5 0 30
Lampyridae 4 21 17 5
Lygaeidae 0 0 0 0
Elateridae 18 5 0 12
Noctuidae 0 0 0 1
Apidae 0 0 1 0
Cynipidae 0 0 18 3
Vespidae 0 0 5 8
Chrysididae 0 0 3 3
Pompilidae 0 0 1 0
Scoliidae 0 0 1 0
Thyreocoridae 0 0 0 14
Berytidae 0 0 0 41
Alydidae 0 0 0 2

 1Unspecified taxa were not identified to the family level.


Table 3. Means (± standard errors) of feeding guilds within farm site and soybean development stage.

Abundance per 10 Sweeps
Feeding Guild Site1 R1 R2 R3 R4 R5
Spider BV 1.22 ± 0.26 a2 0.91 ± 0.20 a 1.27 ± 0.13 a 2.02 ± 0.25 a 1.28 ± 0.22 a
UM 1.19 ± 0.33 a 0.58 ± 0.11 a 0.96 ± 0.10 a 1.19 ± 0.18 a 1.25 ± 0.19 a
Parasitoid BV 0.97 ± 0.20 b 1.28 ± 0.28 ab 1.23 ± 0.15 ab 3.33 ± 0.52 a 4.81 ± 0.92 a
UM 0.22 ± 0.11 b 0.56 ± 0.12 ab 0.72 ± 0.14 ab 0.97 ± 0.16 ab 1.28 ± 0.38 a
Chewing predator BV 0.63 ± 0.33 b 0.13 ± 0.05 ab 0.82 ± 0.15 ab 1.25 ± 0.24 ab 1.56 ± 0.30 a
UM 0.28 ± 0.10 b 0.02 ± 0.02 ab 0.48 ± 0.10 ab 1.59 ± 0.31 ab 1.50 ± 0.31 a
Sucking predator BV 3.75 ± 0.69 b 3.41 ± 0.51 b 4.03 ± 0.37 ab 8.27 ± 0.65 a 6.41 ± 0.73 ab
UM 2.13 ± 0.34 b 2.59 ± 0.35 b 2.03 ± 0.18 ab 7.84 ± 1.05 a 5.91 ± 0.64 ab
Foliage feeder BV 8.78 ± 0.92 b 7.50 ± 0.86 ab 9.62 ± 0.70 a 6.25 ± 0.85 b 4.06 ± 0.86 b
UM 2.81 ± 0.95 b 1.84 ± 0.26 b 5.17 ± 0.41 a 2.89 ± 0.36 ab 2.41 ± 0.52 b
Plant sucking BV 6.81 ± 1.16 ab 5.66 ± 0.61 a 2.73 ± 0.21 ab 3.20 ± 0.43 b 2.22 ± 0.32 b
UM 3.81 ± 0.72 a 8.61 ± 1.01 a 2.97 ± 0.19 a 15.0 ± 3.66 a 8.38 ± 1.77 a
Pod feeder BV 0.81 ± 0.24 a 0.64 ± 0.15 a 1.23 ± 0.16 a 0.50 ± 0.14 a 1.34 ± 0.36 a
UM 1.00 ± 0.21 a 1.73 ± 0.28 a 1.09 ± 0.15 a 1.69 ± 0.29 a 3.50 ± 0.66 a

1BV = Beltsville, UM = Upper Marlboro

2Different letters within individual rows represent significant differences between growth stages.



The objective of this study was to quantify the impact of cover crop termination method and timing on arthropods within soybean foliage. Cover crop termination practices are known to impact arthropods via resulting residues that remain on the soil surface. Thus, it was believed that different cover crop termination methods examined during this study would influence arthropod abundances differently. As expected, delaying the cover crop termination date resulted in significantly greater biomass of residue in late-kill (LK) and flail-mowed (FM) than in early-kill (EK) treatments. Averaged across years, delaying cover crop termination in FM and LK increased barley biomass relative to EK by 2007.5 kg ha−1 (1791 lbs/acre) at Beltsville and 716.8 kg ha−1 (639.5 lbs/acre) at Upper Marlboro. However, arthropod populations within the soybean foliage responded similarly to treatments regardless of plant biomass differences. Instead, arthropod abundances changed according to soybean growth stage. Chemical (LK) and mechanical (FM) termination tactics also had similar effects on arthropod abundances. This suggests that whether cover crops are killed early or late, or chemically or mechanically by mowing, the resulting arthropod community will be similarly impacted. The EK “early” (early April) and LK “late” (at soybean planting from mid to late May) treatments represent some of the most widely used practices for cover crop termination by Mid-Atlantic soybean producers. The results of our study suggest that these two practices are likely to result in similar species and number of foliar arthropods throughout the different soybean growth stages.


Overall, our results indicate that cover crop termination methods that result in greater cover crop biomass will have no effect on insects and spiders within the soybean foliage. However, if delaying cover crop termination results in greater weed suppression without impacting soybean productivity, this practice should nevertheless make soybean systems more resilient to pest pressure and acceptable by soybean producers.


We thank crews at the Upper Marlboro and Beltsville Research and Education Centers for logistics in establishing and completing field trials. This work or publication was supported by Hatch Project No. MD-ENTO-9107/project accession no. 227029 and the Crop Protection and Pest Management (CPPM), Extension Implementation Program (EIP) award number 2017-70006-27171 from the USDA National Institute of Food and Agriculture, and funding from the Maryland Soybean Board.

Should I Be Worried About Mega-Pests?

Peter Coffey, Agriculture Agent Associate
University of Maryland Extension, Carroll County

If you’re a farmer in the United States then you’re acquainted with the corn earworm, Helicoverpa zea. Maybe you know it as cotton bollworm, tomato fruitworm, sorghum headworm, soybean podworm, or maybe you remember when scientists used to call it Heliothis zea. To keep things from getting confusing we’ll just call it H. zea. No matter what you call it, it’s the same tan colored moth with caterpillars we all remember picking out of the tip of an ear of sweet corn as a kid. As you can guess from the variety of names, H. zea feeds on a lot of different crops. It’s actually been documented to eat over 100 different plants, usually the reproductive part of the plant (the fruit/grain/bean part).

Adult corn earworm
Figure 1. Adult Helicoverpa zea moth. Image: Peter Coffey, University of Maryland.

While H. zea is a devastating pest throughout North and South America, it has a bigger badder older brother. In Europe, Africa, Asia, and Australia they also call this moth the cotton bollworm, but it’s a different species, Helicoverpa armigera (we’ll call it H. armigera). In fact, H. armigera is the parent species to our H. zea. Scientists estimate that about 1.5 million years ago some H. armigera moths made their way over to the Americas, and over time they evolved enough that does we consider them to be a different species, H. zea. To put it another way, if dogs evolved from wolves, then H. zea is a bulldog, and H. armigera is the wolf. This is important because H. armigera as the older original species is more genetically diverse, which is probably why it’s better at evolving pesticide resistance.

In recent years, increasing global trade has increased the occurrence of introductions of invasive pests worldwide, and H. armigera has been one of the species that people are the most worried about. It has been caught at the borders several times in North America, but unfortunately in 2013 it was confirmed to be established in Brazil. Additionally, scientists have known for a while that H. armigera and H. zea could mate and create viable hybrids in the laboratory, and just this winter a paper was published documenting the first wild hybrids discovered in Brazil.

Hybrid moths are concerning, because even though H. armigera is the more genetically diverse species, H. zea has spent the last 1.5 million years developing its own unique set of genes. This means that the hybrids combine the genetic diversity of both species. If you’re wondering why genetic diversity is important, remember that when a population lacks genetic diversity we call it inbred, so you could think of hybrids as the opposite of inbred. Scientists worry that these hybrids could attack an even wider range of crops, and that they could evolve pesticide resistance even faster. There’s no reason to think that individual caterpillars would be more damaging, both species are cannibals, which is why you rarely see more than one worm in an ear of corn.

So what does this mean for you? It means that sometime in the future, if these hybrids make it here, pesticide resistance may become even more of a concern than it already is. For now, if you’re spraying corn remember to rotate your chemical families. You can also check out what the corn earworm population is doing at, or call your local extension office and ask.

Protecting Pollinators in Ag Landscapes

Veronica Johnson, Maryland Department of Agriculture

A pollinator is any organism that transfers pollen –the male genetic material of plants- from one flower to the next, resulting in the production of fertile seeds. Pollinators include birds, bats, bees, butterflies, beetles and some small mammals. Bees are the most efficient of the pollinators, with some species capable of visiting up to 6 thousand flowers in a single day. This high rate of flower visitation is important considering between 75% and 95% of all flowering plants on Earth need help with pollination. These plants include the countless fruits, vegetables and nuts that constitute an important part of our diets. In fact, pollinators are responsible for one out of every three bites of food that we eat. Honey bees alone contribute between $1.2 and $5.4 billion in agricultural productivity in the U.S. Unfortunately, pollinator populations are changing. Many populations are in decline, primarily due to loss of feeding and nesting habitats. However, pollution, chemical misuse, disease and changing weather patterns are also contributing to shrinking pollinator populations. Continue reading Protecting Pollinators in Ag Landscapes

Sorghum Growers Encouraged to Keep an Eye Out for Sugarcane Aphid this Season

Kelly Hamby, Assistant Professor/Extension Specialist, Department of Entomology

Ben Beale, Extension Educator, UME-St. Mary’s County

Sugarcane Aphid was found late last fall in Charles County, Maryland in a sorghum field that was being harvested for grain. Aphid populations were very high, with feeding present in the grain head and leaves. This is the first time that sugarcane aphid has been found in Maryland. While this aphid has caused substantial losses to sorghum in states to our South, it is unknown if the aphid will be present early enough and at high enough populations to cause significant injury in Maryland. Growers are encouraged to monitor sorghum fields through the summer for the presence of sugarcane aphid. We suspect sugarcane aphids are most likely to arrive later in the season in Maryland. Continue reading Sorghum Growers Encouraged to Keep an Eye Out for Sugarcane Aphid this Season